Physiological, mitochondrial, and oxidative stress differences in the presence or absence of lactation in rats

Rats were fasted for 4 h and then were anesthetized using isoflurane vapors and body mass was quickly recorded. The anesthetized animals were than decapitated, and blood was collected, allowed to clot, and then centrifuged. Following centrifugation the serum was frozen at −80 °C for subsequent analyses. Serum glucose (STA-680, Cell Biolabs, San Diego, CA, USA) and non-esterified fatty acids (NEFA) (STA-618, Cell Biolabs) were quantified using the manufacturer’s specifications.

After the decapitation, the following tissues were excised and weighed: liver, triceps surae (calf muscle), and the retroperitoneal white adipose tissue (WAT). After the mass of each tissue was recorded, a sample of tissue from calf skeletal muscle and liver was used for mitochondrial isolation and the remainder of tissues were frozen in liquid nitrogen and stored at −80 °C for subsequent analyses. Upon gross evaluation, mammary tissue was not observed for females in any of the groups suggesting that the regression of mammary tissue was nearly or fully complete for females in the PP and PL groups at the time of tissue collection.

Mitochondrial isolations for muscle were performed as previously described [19]. Excised muscles (~750 mg) were trimmed to remove fat and connective tissues, weighed, and placed in 10 volumes of solution I (100 mM KCl, 40 mM Tris HCl, 10 mM Tris base, 1 mM MgSO4, 0.1 mM EDTA, 0.2 mM ATP, and 2% (wt/vol) free fatty acid bovine serum albumin (BSA), pH 7.40). Muscles were minced with scissors and the mince was homogenized for 15 s with a polytron. Protease (Trypsin) was added (5 mg/g wet muscle), and the digested mince was mixed continually for 7 min. Digestion was terminated by the addition of an equal volume of solution I. The homogenate was centrifuged at 500 g for 10 min at 4 °C and the supernatant was rapidly decanted through a double layer of cheesecloth and centrifuged at 3500 g for 10 min at 4 °C. The supernatant was discarded and the mitochondrial pellet was resuspended in solution I. The suspension was centrifuged at 3500 g for 10 min at 4 °C. The supernatant was again discarded, and the pellet was resuspended in 10 volumes of solution II (similar to solution I, but without BSA). This resuspended pellet was subsequently centrifuged at 3500 g for 10 min at 4 °C. The final pellet containing mitochondria was suspended in 250 μl of a solution containing 220 mM mannitol, 70 mM sucrose, 10 mM Tris HCl, and 1 mM EGTA, pH 7.40. Mitochondria from liver were isolated as previously described [20]. Briefly, liver (~750 mg) was weighed and placed in 10 volumes of solution III (250 mM sucrose, 5 mM HEPES, and 1 mM EGTA), minced with scissors and the mince was homogenized with a Potter-Elvehjem PTFE pestle and glass tube (2 passes). The homogenate was centrifuged at 500 g for 10 min at 4 °C and the supernatant was rapidly decanted through a double layer of cheesecloth and centrifuged at 3500 g for 10 min at 4 °C. The supernatant was discarded and the mitochondrial pellet was resuspended in solution III. The suspension was centrifuged at 3500 g for 10 min at 4 °C. The final pellet containing mitochondria was suspended in 250 μl of a solution containing (in mM) 220 mannitol, 70 sucrose, 10 Tris HCl, and 1 EGTA, pH 7.40. All procedures we performed on ice and all solutions were kept on ice (4 °C).

Mitochondrial oxygen consumption was measured as described by Messer et al. [21]. Briefly, mitochondrial oxygen consumption was measured polarographically in a respiration chamber (Hansatech Instruments, United Kingdom). Isolated mitochondria (20 μL) were incubated with 1 ml of respiration buffer adapted from Wanders et al. [22] (100 mM KCL, 50 mM MOPS, 10 mM KH₂PO₄, 20 mM glucose, 10 mM MgCl₂, 1 mM EGTA, and 0.2% fatty acid free BSA; pH = 7.0) at 37 °C in a respiratory chamber with continuous stirring. For state 3 respiration, 2 mM pyruvate and 2 mM malate (complex I substrates) or 5 mM succinate (complex II substrate) was used in the presence of 0.25 mM ADP, and state 4 respiration was recorded following the phosphorylation of ADP as described by Estabrook [23]. Respiratory control ratio (RCR) was calculated as state 3/state 4 oxygen consumption. Respiration values were expressed as a ratio to citrate synthase to compensate for mitochondrial enrichment in the samples.

Oxidant emission by mitochondria was determined using the oxidation of the fluorogenic indicator Amplex Red (Molecular Probes, Eugene, OR) in the presence of horseradish peroxidase [24]. The assay was performed at 37 °C in 96-well plates using succinate as the substrate. Specifically, this assay was developed based on the concept that horseradish peroxidase catalyzes the hydrogen peroxide-dependent oxidation of nonfluorescent Amplex Red to fluorescent Resorufin Red. Resorufin Red formation was monitored at an excitation wavelength of 545 nm and an emission wavelength of 590 nm using a multiwell plate reader fluorometer (Synergy H1, BioTek, Winooski, VT, USA). We recorded the level of Resorufin Red formation, and hydrogen peroxide production was calculated with a standard curve.

Complex I (NADH dehydrogenase) enzyme activity (EC 1.6.5.3) was measured as a function of the decrease in absorbance from NADH oxidation by decylubiquinone before and after rotenone addition [25]. Complex II (succinate dehydrogenase) activity (EC 1.3.5.1) was measured as a function of the decrease in absorbance from 2,6-dichloroindophenol reduction [25]. Complex III (ubiquinol cytochrome c oxidoreductase) activity (EC 1.10.2.2) was determined as a function of the increase in absorbance from cytochrome c reduction [25]. Complex IV (cytochrome c oxidoreductase) activity was determined as a function of the decrease in absorbance from cytochrome c oxidation [25]. Specificity of complex IV activity was determined by monitoring changes in absorbance in the presence of KCN [25]. Citrate synthase (EC 4.1.3.7) was measured as a function of the increase in absorbance from 5,5′-dithiobis-2-nitrobenzoic acid reduction [25]. Enzyme activities were expressed as a ratio to citrate synthase to compensate for mitochondrial enrichment in the cell samples. All measurements were performed using the BioTek Synergy H1 spectrophotometer (BioTek, Winooski, VT, USA).

The relative concentration of proteins was quantified by Western blot analysis [24]. To accomplish this, tissue was homogenized 1:10 (wt/vol) in 5 mM Tris HCl (pH 7.5) and 5 mM EDTA (pH 8.0), and protease inhibitor cocktail (14224–396, VWR, Radnor, PA, USA) and was centrifuged at 1500 g for 10 min at 4 °C. Protein content of the supernatant was quantified by the method of Bradford [26]. Proteins were separated by polyacrylamide gel electrophoresis via 4–20% polyacrylamide gels (BioRad, Hercules, CA, USA). After electrophoresis, the proteins were transferred to PVDF membranes. Non-specific sites were blocked in phosphate-buffered saline (PBS) solution containing 0.1% Tween 20 and 5% non-fat milk. Membranes were then incubated overnight at 4 °C with primary antibodies purchased from GeneTex (Irvine, CA, USA) directed against peroxisome proliferator activated receptor alpha (PPARα, GTX101096, 1:1000), peroxisome proliferator activated receptor delta (PPARδ, GTX113250, 1:2000), peroxisome proliferator activated receptor gamma, coactivator 1 alpha (PGC-1α, GTX37356, 1:1000), superoxide dismutase 1 (SOD1, GTX100554 1:2000), superoxide dismutase 2 (SOD2, GTX116093, 1:2000), catalase (CAT, GTX110704, 1:2000), and glutathione peroxidase (GPX, GTX116040, 1:2000). Following incubation with primary antibodies, membranes were washed with PBS-Tween (5 min × 3) and then incubated with secondary antibodies for 1 h in room temperature. After washing (5 min × 3), a chemiluminescent system was used to detect labeled proteins (GE Healthcare, Buckinghamshire, UK). Images of the membranes were captured and analyzed by using the ChemiDoc-It2 Imaging System (UVP, LLC, Upland, CA). Protein expression was normalized to Ponceau staining.

To determine the relative amount oxidative damage, we measured protein oxidation and lipid peroxidation. Lipid peroxidation was assessed by determining 4-hydroxynoneal (4-HNE; trans-4-hydroxy-2-nonenal, C₉H₁₆O₂) expression via western blotting, as described above. Primary antibody for 4-HNE was purchased from Abcam (ab46545; 1:1000 dilution, Cambridge, MA, USA). Protein oxidation was measured by comparing relative expression of protein carbonyls using a commercially available kit (Oxy-Blot protein oxidation detection kit; Intergen, Purchase, NY, USA) via western blotting, as described by the manufacturer’s instructions.

During pregnancy and lactation, the body of females undergoes a large fluctuation in mass to accommodate the developing fetuses and active mammary tissue. The regression of these tissues 28 days following pregnancy and lactation varied between groups. Body mass was higher in PP compared to NR rats (p = 0.005) and the body mass of PP rats was higher than the mass of the PL rats, but this was not statistically different (p = 0.145) (Fig. 1a). Liver mass was higher in PL than NR rats (p = 0.004) (Fig. 1b). PL rats had lower retroperitoneal fat mass compared to both PP (p = 0.041) and NR rats (p < 0.001) (Fig. 1c). The combined mass of the rear triceps surae calf muscle was higher in PP than in NR rats (p = 0.027) (Fig. 1d).

The high energetic demand of lactation requires energy substrates to be delivered to metabolically active tissues via the blood stream. Two important metabolites transported in blood are glucose and NEFAs. Serum glucose concentration was lower in PL compared to NR (p = 0.029), but no statistical differences were detected in serum NEFA concentrations (p = 0.802) (Fig. 1e and f).

RCR is a useful tool in assessing mitochondrial function, and can be used to measure respiration capacity of the mitochondria with different available substrates. Measuring RCR using isolated mitochondria provides the ability to observe ATP phosphorylation via electron shuttling at complex I and complex II. High-resolution respirometry was used in isolated mitochondria from liver and skeletal muscle. The RCR of liver mitochondria was higher in PL compared to PP and NR rats (p = 0.029 and p = 0.002, respectively) when using pyruvate and malate as the substrate (Fig. 2c). No statistical differences were detected in liver RCR between groups when succinate was used as the substrate (p = 0.234; Fig. 2f). In addition, no statistical differences were detected in skeletal muscle RCR between groups when pyruvate and malate or succinate was used as the substrate (p = 0.792 and p = 0.996, respectively) (Fig. 2i and l). Oxidant emission occurs during oxidative phosphorylation in the mitochondria, primarily at complex I and complex III sites [27]. No statistically differences (p = 0.191) were detected in oxidant emission from mitochondria isolated from liver (Fig. 2m). Oxidant emission from mitochondria isolated from skeletal muscle was lower in PP and PL compared to NR (p = 0.017 and p = 0.008, respectively) (Fig. 2n).

Phosphorylating ADP in the mitochondria is accomplished through electron pumps that harness the energy of electrons by oxidizing the reduced form of either NADH or FADH. Specifically, complex I transfers electrons from NADH to coenzyme Q10. Complex II transfers electrons from FADH. Complex III passes electrons down the chain by reducing cytochrome c. And, complex IV converts molecular oxygen into water and further shuttles protons into the intermembrane space. Activity of complex I in liver mitochondria was higher for both PP and PL rats compared to NR rats (p = 0.027 and p = 0.046, respectively) (Fig. 3a). Conversely, enzymatic activity of complex II in liver mitochondria was lower in PL rats than both NR and PP rats (p = 0.004 and p = 0.0029, respectively) (Fig. 3b). Complex III activity in liver mitochondria of PL rats was lower, albeit not statistically different, than PP and NR rats (p = 0.065 and p = 0.075, respectively) (Fig. 3b). Complex IV activity was lower in liver mitochondria of PL compared to PP rats (p = 0.025; Fig. 3d). Enzymatic activity of complex I and II were lower in mitochondria isolated from skeletal muscle in PL compared to NR rats (p = 0.044 and p = 0.032, respectively). (Fig. 3e and f).

The PPAR superfamily is associated with the regulation of genes involved in oxidative metabolism. Specifically PGC-1α is associated with regulation of genes involved in mitochondrial biogenesis [28] and PPARδ is associated with genes involved in lipid and glucose metabolism [29, 30]. PGC-1α liver protein expression was higher in PL rats compared to PP and NR rats (p = 0.035 and p = 0.001, respectively) (Fig. 4a). Also, PPARδ liver protein expression was higher in PL compared to NR rats (p = 0.009) (Fig. 4b). No statistically differences were detected for PGC-1α and PPARδ in skeletal muscle (p = 0.331 and p = 0.691, respectively) (Fig. 4c and d). PPARδ protein expression in WAT was higher in PL compared to PP and NR rats (p < 0.001) (Fig. 4e).

The balance between oxidants and antioxidants is often referred to as oxidative stress. Endogenous antioxidant defense mechanisms exist to protect and detoxify oxidants. Specifically, SOD2 and SOD1 act to detoxify superoxide into the lesser harmful hydrogen peroxide. Hydrogen peroxide can be then be further detoxified by CAT and GPX. The protein levels of the antioxidants (SOD2, SOD1, CAT, and GPX) and markers of oxidative damage were compared between groups. Liver SOD2 protein levels were higher in PL compared to NR and PP rats (p = 0.001 and p = 0.025, respectively) (Fig. 5a). Liver CAT protein expression was higher in PL and PP rats compared to NR rats (p = 0.015 and p = 0.001, respectively) (Fig. 5c). No statistical differences (p > 0.05) were detected for SOD1, GPX, 4-HNE or protein carbonyls in liver (Fig. 5b and d-f). SOD2 protein levels were higher in WAT of PL compared to NR rats (p = 0.039) (Fig. 6a), and 4-HNE was lower in WAT of PL rats compared to NR rats (p = 0.004) (Fig. 6e). No statistical differences (p > 0.05) were detected for SOD2, SOD1, CAT, GPX, 4-HNE, or protein carbonyls in skeletal muscle (Fig. 7).

The process of milk synthesis presents a high energetic demand to the mother. In mice, resting metabolic rate during peak lactation has been recorded as being 2-fold higher than controls [33]. A large contribution to this increase in resting metabolic rate is imposed by the liver, which doubles in size, and the mammary gland that is actively synthesizing milk. Our findings show that PP rats had a higher body mass 28 days post-partum compared to NR, while no statistical differences were detected between PL and NR rats. Furthermore, our results indicate that while body mass was not different between NR and PL rats, WAT mass was lower in PL rats compared to both PP and NR rats. These finding agree with previous work showing lactation leads to a reduction in absolute fat mass associated with fewer visceral adipocytes [34‐36]. In addition, liver is hypertrophied during gestation and lactation in order to meet the demands of higher energy utilization. Our data support this, and liver mass was still higher in PL rats 7 days after weaning, which may have masked some of the difference in body mass between the groups. These findings highlight the extent of anatomical changes that occur in response to the metabolic perturbations imposed by lactation.

Systemic changes in numerous metabolic processes must occur during pregnancy for the female’s body to support the growing fetus. One such change is an increase in maternal insulin production. An increase in maternal insulin production facilitates glucose transport to the fetus, but this can often result in increased insulin resistance in maternal cells. In some cases, this may manifest itself as gestational diabetes. We show that alterations of glucose metabolism due to pregnancy is sustained to some degree in females that do not participate in lactation. This idea is supported through epidemiological findings suggesting that this effect increases a woman’s risk of developing type II diabetes later in life relative to women who do breastfeed [5, 37]. In contrast to pregnancy, there is a decrease in insulin secretion during lactation returning to pre-pregnancy levels that is associated with a decline in β-cell proliferation and improved insulin sensitivity [13].

Importantly, our finding of higher liver protein levels of PPARδ in PL rats may help explain how glucose metabolism may be increased due to a lactation period. PPARδ plays a significant role in the regulation of glucose metabolism and insulin sensitivity [29] and Sanderson and collaborators demonstrated its importance in the liver. Specifically, knockout models of PPARδ had lower expression of genes relating to glucose metabolism and higher plasma glucose in a fasted state compared to wild type mice [38]. These studies and our data, in which fasted serum glucose was decreased and liver PPARδ protein expression was higher in PL rats, suggest that the resulting modulation of PPARδ expression persists after lactation and is at least one mechanism that may confer a phenotype that protects against type II diabetes. In addition, we report that mitochondrial respiratory function was enhanced in PL rats when using complex I substrates, as indicated by a higher RCR. Considering that complex I respiration plays a large role in the utilization of glucose, via high NADH:FADH₂ production, our findings provides a mechanistic outcome for the enhanced ability of cells in PL rats to metabolize glucose. Interestingly, measurements of enzymatic activity of the electron transport system show that both PP and PL rats had higher complex I activity. Our findings of increased liver complex I mitochondrial function in PL animals may be better explained by the increased protein expression of PGC-1α and PPARδ. While PPARδ knockout models have demonstrated that PPARδ is involved in the expression of genes relating to oxidative metabolism, PGC-1α is a well-known regulator of mitochondrial biogenesis and coactivator of genes involved in oxidative phosphorylation [28]. Thus, increased expression of these proteins likely results in the improved mitochondrial function observed. PL rats also expressed lower liver complex II activity compared to both PP and NR, and lower respiration when using complex II substrates. This down regulation of complex II respiration further limits use of FADH₂ to fuel OXPHOS, and as a result, likely down regulates fat metabolism via beta oxidation, an effect that further supports the use of glucose as the primary fuel to support ATP production in PL rats. During pregnancy, the mother’s body experiences increasing visceral adiposity [12]. During lactation however, a shift in lipoprotein lipase and triacylglyceride levels facilitate the mobilization of fats to the mammary gland to be used for milk synthesis [14]. In contrast to PPARδ’s role in liver, PPARδ in WAT is involved in catabolic effects through oxidation of fatty acids [30]. Our data show that PPARδ protein expression is higher in WAT of PL rats compared to NR and PP rats. Our findings of increased PPARδ protein expression in two functionally different tissues also demonstrate that a systemic signaling event is a likely candidate for driving these changes.

The production of reactive oxygen species (ROS) is a naturally occurring phenomenon that can manifest itself through processes of energy production and metabolism; however an imbalance between ROS production and a counteracting antioxidant system can result in oxidative damage [39]. Oxidative stress has been implicated as a driving force in the development of diabetes and other health disparities [40, 41] and several investigators have theorized that oxidative stress is a physiological cost of reproduction that has the potential to impact a female’s future reproductive performance and longevity [42, 43]. Yet, the results of studies testing this hypothesis have provided equivocal results [18]. Our data show no differences in markers of liver lipid peroxidation and protein oxidation between the groups described here that were collected 1 week after lactation has ended, suggesting that there was no persistent oxidative cost of reproduction. With no difference in oxidant emission in liver, our finding of higher liver SOD2 in PL rats may be better explained by increased PGC-1α, as SOD2 is a downstream target of PGC-1α [44] than as a mechanism of preventing oxidative damage. Systemic effects of prior lactation are further demonstrated by our finding that SOD2 protein levels were also higher in the WAT of PL rats. However, oxidative damage was lower in WAT of PL as shown by decreased 4-HNE. Reduced oxidative damage in WAT following lactation may serve as a beneficial adaptation, as oxidative stress in WAT has been proposed to disrupt adipokine secretion by WAT, which may play a role in the development of metabolic syndrome [41].